Field sampling procedure

Prior to biophysical sampling at each station, spatial reference (latitude and longitude) along with prevailing weather conditions were recorded in a field logbook. The near-bottom environmental variables (conductivity (mS/cm), temperature (°C) and depth (m)) were recorded by means of a single multi-parameter profiler CTD (Sea-Bird: SBE 19 Plus v.2) deployed to just above the seafloor (deployment speed 1 m s^–1), equipped with auxiliary sensors to measure additional parameters. These included concentration of dissolved oxygen (mg.l^-1), pH, turbidity (NTU) and fluorescence content (mg.l^-1) (McArthur et al. 2010a, MacKay 2014). Only the bottom 5 m hydrographic characteristics measured at each station were used for further analysis.

Grab sampling commenced once the CTD was retrieved on deck.

Figure 2.2. KZN biodiversity planning Biozones and the associated sampling stations. These are depicted relative to bathymetry (50-80 m depth shown as red lines) and sampling stations.

2.3.2 Macrobenthic and sediment sampling

Sample collection was limited to daytime hours (MacKay 2014), when many mobile species can be expected to be relatively inactive. Prior to grab deployment, a drop video was cast to the seafloor to validate the extent of unconsolidated sediments to avoid sampling reef (MacKay 2014). Triplicate biological samples were collected from each station, using a long armed 0.2 m² van Veen grab and extra weights to 40 kg. A minimum of three replicates were collected to increase the probability of sampling rare taxa (in this context refer to individuals with low abundance or small range size (Ellingsen 2001)) and are considered appropriate for statistical comparison (Gray and Elliott 2009). The grab was deployed through a mechanical winch over the side of the vessel (MacKay 2014).

Bucket type grabs such as the van Veen are appropriate in soft-bottoms, although the volumes of sediment sampled are mostly affected by the type of substrate, with samples collected from muddy bottoms often filling the grab completely, while samples collected in sand to gravel substrates often penetrate to minimum depth (Somerfield et al. 2005, Tagliapietra and Sigovini 2010). Care was taken to ensure all replicates were within 100 m of each other. Once a grabbed sample was retrieved, sediment depth was measured to the nearest mm through a top opening window on the grab. A sample was acceptable and considered a successful grab, only if vertical sediment depth was at least 50 mm or more with no indication of disturbance (MacKay 2014).

The interrelation between macrobenthos and sediments requires characterisation of the bottom environment. Therefore, a sub-sample (175 g) was taken from the upper 5 cm from each grabbed sample for bicarbonate content, sand grain size and organic content analysis. The latter required fixation by addition of 40% formalin (MacKay 2014). The remaining biological sample was washed through a square stainless steel sieve with 1000 µm mesh size. Square mesh sieves are advisable as they provide a larger percentage of open area (Somerfield et al. 2005).

Sediment colour and the odour were noted in a field trip logbook before washing. Any visible fauna were carefully handpicked and bottled in the suitably labelled sample jar. Sieving was gentle to minimise sediment being displaced out of the sieve and the impact of strong water pressure on delicate animals (MacKay 2014). Material remaining on the mesh sieve after washing, consisting of benthic animals, tubes, shells, shell hash, and coarse sediments, were bottled in a suitable biological sample jar and preserved unstained with buffered forma-saline (4%) allowing for subsequent analysis ashore (MacKay 2014).

GENERAL MATERIAL AND METHODS

MACROBENTHOS AND MARINE BIODIVERSITY SPATIAL ZONES

26 In certain cases samples (e.g. at stations S1 and S3) were dominated by coarse sediments (sand and gravel >10 kg) that were retained on the sieve and precluded the sorting of infauna in a reasonable timeframe. In these cases, elutriation was conducted to separate light material and small animals from sediments (Wilson 2005, MacKay 2014). The process: 2 litres of unsorted sediments from the larger sample was put into a 20 L bucket three-quarter filled with seawater and well agitated to suspend light material and small animals. Floating animals and materials were collected by pouring the agitated volume of water through a 1000 µm sieve before resettlement. This was repeated at least five times or until the water ran clear before sorting for large and heavy fauna such as molluscs and characteristic sedimentary elements (Somerfield et al. 2005, Wilson 2005, MacKay 2014).

2.3.3 Laboratory procedure Sedimentary analysis

The sedimentary analysis was performed at a sedimentological analysis laboratory (Environmental Mapping and Survey (EMS)) on samples collected from every grab. The granulometric analysis was used to characterise the sedimentary environment and provides an insight into local physical conditions.

Grain size analysis

The aim of grain size analysis is to define relative proportions of different grain sizes which make up a given sediment population (Gray 1981, Bale and Kenny 2005). A number of techniques have been defined for grain size analysis (Gray and Elliott 2009). In this regard, a sieving technique was employed as a result of the advantages associated with it, such as being less expensive, an easily reproducible method and deemed the most practical way of characterising particle size larger than 0.063 mm (Bale and Kenny 2005).

Grade scales apply random set of finite ranges to the continuous frequency distribution of particle sizes in order to produce logical classifications for the numerical divisions (Gray 1981, Bale and Kenny 2005). The Wentworth (1922) ordination scale is most commonly used by marine ecologists and geologists to characterise sediment particles (Bale and Kenny 2005, Gray and Elliott 2009). By applying a logarithmic scale transformation (millimetres into whole integers) to the Wentworth scale produces the phi (Φ) notation which was initially employed to graphically manipulate data (Gray 1981, Hayes et al. 1992, Bale and Kenny 2005), based on the following definition: phi units (F) = -log2 (diameter in mm). This was done to graphically manipulate data (Gray 1981, Bale and Kenny 2005). Three general categories (gravel, sand, and mud), classified according to the dominant size of the individual clasts are recognised.For data

analysis in this study, the substratum was retrospectively subdivided into seven categories (Table 2.1).

Sorting

Grain size alone does not account for particle size variation or the particle shape, therefore applicability of this measure is limited in describing a highly unimodal sediment grain size distributions (Lewis and McConchie 1994). Sorting is another measure used to characterise grain size according to sorting classes (Table 2.2). Sorting, or grain-size variation reflects the amount of interstitial space available within sediment as a habitat for biota (Gray 1974b). Well-sorted sediments are generally associated with a high energy or dynamic environment and the majority of the sample comprised of particles with the same diameter (Mackay 2006, Gray and Elliott 2009). In contrast, poorly-sorted sediments mostly comprise various grain sizes and are mostly associated with a stable environment, less frequently affected by hydrodynamics (Gray and Elliott 2009, McArthur et al. 2010a).

Skewness

This descriptive measure indicates symmetry, which is a preferential distribution to one side of the mean (Gray and Elliott 2009). Symmetrical sediments will have a value of 0.00, whereas those with more courser grain-size are positively skewed (- phi values). Negative skewed (+ phi values) sediments are dominated by finer grained size (Table 2.3) (Gray 1981).

Organic content

Features of organic matter include their ability to form water-soluble and insoluble complexes, interact with clay minerals, bind particles together, to absorb and release organic compounds

Table 2.1. Wentworth scale of sediment particle size (mm) classification with the accompanying phi (ɸ) notation. (Adopted from Wentworth 1922 and Gray 1981).

Sediment type Grain size (mm) Phi (ɸ) scale

Gravel > 2 < -1.0 Silt and clay fraction < 0.0625 > 4.0

Table 2.2. Sediment sorting classes used to classify sediments (modified from Gray 1981).

phi (ɸ) scale

GENERAL MATERIAL AND METHODS

MACROBENTHOS AND MARINE BIODIVERSITY SPATIAL ZONES

28 and plant nutrients and retain water in the sediment (Schumacher 2002). Therefore, examination of total organic carbon is critical, since organic materials present in sediments can be expressed as carbon, and carbon is available in inorganic and organic form, the latter being an important food resource for macrobenthic communities (Sverdrup et al. 1942, Cocito et al. 1990, Méndez and Green 1998)

A suite of techniques have been developed to determine organic content, all relying on the principle of destroying organic matter present in a given set of sediments through chemical or heat energy and then measuring the loss directly (Schumacher 2002). This study used a chemically-based digestion technique to determine total organic content (TOC) of sediments.

This was adopted from Schumacher (2002) and the basic steps for this technique are:

1. Inorganic carbonates are eliminated by adding 6% Hydrogen peroxide (H2O2) to a known weight of sediment until the frothing reaction ceases.

2. The sample is incinerated at 105°C, cooled and weighed.

3. Organic matter present in the sediment sample is determined gravimetrically and calculated as:

𝑂𝑀 =^{𝑊i−𝑊f}

𝑊i × 100%

(2.1)

Where OM is organic matter (%), Wi is the initial sample weight (g) and Wf the final sample weight (g).

4. Organic matter is converted to a value of TOC (Table 2.4) by using an appropriate factor. A conversion factor of 1.72 is commonly used, based on the assumption that organic matter contains 58-60% Carbon (Schumacher 2002).

Table 2.3. Skewness categories used to indicate a proportion of sediments grain-size (adopted from Folk 1974).

Verbal classification

from to Graphically skewed to

1 0.3 Strongly + skewed Very – phi values (coarse sand)

0.3 0.1 + skewed - phi values

0.1 -0.1 Nearly symmetrical symmetrical

-0.1 -0.3 - Skewed + phi value

-0.3 -1 Strongly - skewed Very + phi values (fine sand) Values

symmetry

Macrobenthos

In the laboratory, prior to sorting, samples were rinsed with seawater through a 1000 µm sieve to remove all traces of formaldehyde. Animals from each sample were separated from non-biogenic material under a magnifying lamp/microscope and sorted according to major taxonomic groups (polychaetes, bivalves, decapods, amphipods and isopods) to aid taxonomic identification. To reduce the risk of human error, sorted samples were cross-checked between sorters to ensure that no animals were left. Sorted fauna were then identified to coarse operational taxonomic units (OTUs) by a single researcher to minimise observer bias.

Stereomicroscopes (Zeiss stemi-DV4, Zeiss Stereo v.12) and appropriate taxonomic keys (Clark 1923, Barnard 1950, Day 1967, Kensley 1972, Griffiths 1976, Kensley 1978, Kilburn and Rippey 1982) and online keys (e.g. Crustacea.net (Lowry 1999)) were used for taxonomic identification. The species level (lower taxonomic resolution) is the level at which animals interact with their habitat (Bertrand et al. 2006), and quantification of biodiversity and the utility of potential surrogates therefore are often affected by the taxonomic resolution (Bertrand et al.

2006); coarse taxonomic resolution providing rudimentary spatial variation (Anderson et al.

2005).

In the case of a damaged specimen or when some of the diagnostic morphological features were not observable, a staining procedure (Methyl blue) was used (Winsnes 1985). Taxonomic identification was checked and verified by a macrobenthic scientist (C.F. MacKay at Oceanographic Research Institute). To confirm a species name, taxonomic information was checked against the World Registry of Marine Species (WoRMS Editorial Board n.d.) database and the nomenclature herein was based on the latest available taxonomic information during the drafting of this thesis. The number of individuals of species was counted and recorded together with sample details (e.g. date and sample location). Larval and pelagic fauna were excluded and only benthic fauna were considered for further analysis.